An analysis of methods for permanently mounting ovules cleared in four-and-a-half type clearing fluids

Ovules cleared in benzyl benzoate-4 1/2 clearing fluid can be permanently mounted in Piccolyte or Permount by replacing the cleaning fluid with absolute ethanol, upgrading the ovules in mixtures of ethanol and xylene (3:1, 2:2, 1:3, and xylene), and mounting them in either mountant under the supported coverglass of a Raj slide. Optical saggittal sections through the ovules resemble microtome sections in that the protoplasts are slightly shrunken away from the cell walls. The artifact is common in permanently mounted sections; fixation and paraffin infiltration are usually cited as the causes--its appearance in the whole-mounted ovules is caused by xylene. Although miscible with the clearing fluid, Euparal is the least satisfactory of the standard mountants for permanent preparations of cleared ovules and is best used with an equal quantity of clearing fluid for semipermanent preparations. A large quantity of Euparal in the mountant produces pronounced shrinkage. A method for permanently mounting cleared ovules with the clearing image unaltered employs a mountant which contains the ingredients of Spurr low viscosity embedding medium. Vinylcyclohexene dioxide (10 drops) is combined with diglycidyl ether of polypropylglycol (6 drops) and nonenyl succinic anhydride (26 drops). Ovules treated for 24 hr in benzyl benzoate-4 1/2 clearing fluid are passed through a graded series of clearing fluid-epoxy medium mixtures (3:1, 2:2, 1:3, and pure epoxy medium) at intervals of 14 minutes. One drop of dimethylaminoethanol, the cure accelerator, is then added to the epoxy medium and the ovules are mounted and covered immediately on a Raj slide. The preparation is cured in an oven at 60 C for 24 hr and observed with phase contrast or Nomarski interference optics.     

A Trichrome Stain for Insect Material

Deparaffinized insect sections are brought down to water and overstained in a 0.1% solution of azocarmine G in 1% acetic acid. They are then destained in a saturated solution of orange G until the azocarmine G is removed from the endocuticle and the latter is colored pale yellow. After washing, the sections are transferred to a 5.0 % solution of phosphotungstic acid in water for 3 min. They are then rinsed in distilled water and stained in a 0.1% solution of methyl green in 1% acetic acid until the endocuticle is green. Differentiation is done in 2 changes of 95% alcohol. The sections are then dehydrated either in absolute alcohol or dioxane, cleared in a mixture of “camsal”, eucalyptol, dioxane, and paraldehyde (1:2:2:1), and mounted in Mohr and Wehrle's medium, a mountant of the Euparal type.     

Use of the Four-and-a-Half Clearing Technique to Study Gymnosperm EmbryoIogy: Cunninghamia lanceolata

Since its introduction in 1971, the four-and-a-half clearing technique has been widely applied to the study of ovule and female gametophyte development in flowering plants as an alternative to the more arduous paraffin section methods. The technique has undergone several modifications that have broadened its application in studies of Angiosperm embryology. To date, however, the technique has not been successfully applied to embryological features of Gymnosperms. Dark coloration caused by naturally occurring substances and by-products of fixation render the clearing fluid ineffective, and special pretreatment methods used to remove dark substances in Angiosperm ovules have little or no effect on Gymnosperm material. In the technique reported here, paraffin sections of ovules and young seeds of Cunninghamia lanceolata 80-120 μm thick are cleared in benzyl benzoate-412 clearing fluid and examined with phase contrast optics. Observations of the mature female gametophyte in these cleared preparations are compared with those obtained from 10 μm sections, stained with safranin and fast green, and examined with bright-field optics. Although contrast and definition are more pronounced in stained sections than in cleared ones, the differences would not alter one's interpretation of characteristic structural features. The thick, cleared section offers an advantage over the thin, stained one in that many structural entities are contained within a single section rather than spread through several serial sections. The time required for clearing thick sections is much shorter than that required for making permanent stained preparations.     

A Trichrome Stain for Insect Material

Deparaffinized insect sections are brought down to water and overstained in a 0.1% solution of azocarmine G in 1% acetic acid. They are then destained in a saturated solution of orange G until the azocarmine G is removed from the endocuticle and the latter is colored pale yellow. After washing, the sections are transferred to a 5.0 % solution of phosphotungstic acid in water for 3 min. They are then rinsed in distilled water and stained in a 0.1% solution of methyl green in 1% acetic acid until the endocuticle is green. Differentiation is done in 2 changes of 95% alcohol. The sections are then dehydrated either in absolute alcohol or dioxane, cleared in a mixture of “camsal”, eucalyptol, dioxane, and paraldehyde (1:2:2:1), and mounted in Mohr and Wehrle's medium, a mountant of the Euparal type.     

False Cellular Localization of Aminopeptidase Caused by Resinous Mounting Media

Fresh frozen sections of rat submandibular gland were processed for leucine aminopeptidase localization with L-lencyl-β-naphthylamide hydrochloride and L-leucyl-β-naphthylamide. When the first substrate was utilized stained sections were dehydrated in an ethyl alcohol series, cleared in xylene, and mounted in water-insoluble (resinous) media. Sections were also removed at each stage of dehydration and cleared and mounted with glychrogel. When the second substrate was used, tissues were partially decolorized in 40% ethyl alcohol and mounted directly in glychrogel. Comparison of all sections mounted in glychrogel indicated that there was no variation in cellular localization, regardless of substrate used or degree of dehydration. Nuclei were unstained. After mounting in balsam or synthetic resin the nuclei exhibited an intense stain and the parenchymal reaction was stronger, but diffuse. Progressive staining of the nuclei was observed, microscopically, immediately after applying the resinous mounting medium. The use of an aqueous mounting medium appears to be mandatory in this procedure—glychrogel is recommended.     

A NEW CLEARING-SQUASH TECHNIQUE FOR THE STUDY OF OVULE DEVELOPMENT IN ANGIOSPERMS

The simple, efficient method described here for the study of ovule and megagametophyte development in angiosperms provides for the extension of investigation beyond the limits imposed by the traditional but arduous section technique. Excised pistils previously fixed in FPA₅₀ and stored in 70 % ethanol are placed in a clearing fluid composed of lactic acid (85 %), chloral hydrate, phenol, clove oil, and xylene (2:2:2:2:1, by weight). After 24 hr, ovules dissected from the ovularies are transferred with some of the fluid to a slide, covered so that the cover glass is supported laterally by two permanently affixed covers, and examined with phase contrast optics. The unique action of the clearing fluid permits the study of cellular structure with the phase oil objective focused at any focal plane within the ovule. Downward focusing thus reveals a series of optical sections in the sagittal, frontal, or transverse plane depending on the orientation of the ovule. Orientation can be altered by a slight shifting of the cover glass on the lateral support mounts. The ovules become quite fragile in the clearing fluid. Pressure applied to the cover glass gradually breaks the ovule apart without disrupting the structural integrity of individual cells. This squash procedure provides for extending observations to cytological features of megasporocytes, megaspores, and megagametophytes previously identified in intact ovules. The new method is applied here to the study of ovule development in two unrelated species, Cassia abbreviata Oliver var. granitica Bak. f. (Leguminosae) and Ludwigia uruguayensis (Camb.) Hara. (Onagraceae). For best results, the ovules of Ludwigia must be pretreated in lactic acid (85 %) for 24 hr prior to application of the clearing fluid. Other methods for pretreatment likely will be required as the technique is applied to a wider range of flowering plant species.     

Squash Method for Meiosis in Ovules

The ordinary Feulgen or acetic-lacmoid squash tech-nic following fixation in freshly made Carnoy's fluid (alcohol, 6: chloroform, 3: glacial acetic acid, 1), provides an easy and reliable method of studying meiosis in ovules. After fixation for 1 day, the material was hardened in 95% ethyl alcohol for 1-2 days and taken to water by gradual hydration. For staining by the Feulgen method, the material was hydrolyzed 8-10 minutes in 1 N HO at 58-60°C., followed by staining in decolorized leuco basic fuchsin for 2 hours. The staining was intensified by transferring the material to water. After 15-20 minutes the water was replaced by 45% acetic acid. For staining by acetic-lacmoid, the ovules after fixation, hardening and hydration were transferred to standard acetic-lacmoid stain to which was added 1 drop of 1 N HCl to every 10 drops of stain. Gentle heat was applied till the stain started to give fumes. After allowing 20 minutes at room temperature the material was transferred to fresh acetic-lacmoid. Some 6-12 ovules were mounted either in a drop of 45% acetic acid or acetic-lacmoid, depending upon the Feulgen or acetic-lacmoid staining respectively. Gentle and repeated tapping over the cover glass by a blunt needle loosened the cells of integument and nucellus and finally left the megaspore mother cells undergoing meiosis, fully exposed to view. The process was carried out under constant observation using the low power of the microscope. The desired amount of flattening was brought about by light pressure over the cover glass and gentle heating. The preparations were made permanent by dehydrating in ethyl alcohol and mounting in Euparal.     

Fluorescence in Paraffin Sections of Dye-Labelled Protein Administered in vivo: Effect of HgCl2 Fixation

Lung and liver slices, 2-3 mm thick, from guinea pigs injected intravenously with fluorescent dye-protein conjugate are fixed for 15-30 min in saturated aqueous HgCl₂, dehydrated in ethanol, cleared in xylene and embedded in paraffin at 60 C. Mercurial deposits are removed with I₂KI from 5 μ sections taken to water, and the iodine then removed with 5% Na₂S₂O₃. Sections are mounted from xylene into permanent nonfluorescent mounting medium. This procedure gives optimal fluorescence which is not decreased by the technic of removing mercurial precipitates. Longer fixation, fixation in phosphate-buffered formalin, or in an HgCl₂-formalin mixture gives inferior results.     

Non-aqueous permanent mounting for immunofluorescence microscopy

It is generally assumed that an aqueous mounting medium is necessary for the preservation of immunofluorescent-labelled microscopical preparations and polyvinyl alcohol-based solutions (e.g. Mowiol) being the most frequently used mounting media; however, both the quality and intensity of the fluorescence signal in most immunolabelled preparations after aqueous mounting slowly diminish with time, and finally, samples become unsuitable for examination. In the present work, we describe a very simple and rapid non-aqueous mounting procedure for cultured cells and tissue sections, which preserves the fluorescent signal in an excellent way after immunodetection or use of other specific labelling methods. It is based on the current histological protocol in which, after fluorescence labelling, preparations are dehydrated in ethanol, cleared in xylene and mounted in DePeX. Using this non-aqueous mounting medium, the fluorescent signal remains high and stable, allowing a suitable and permanent preservation of labelled and counterstained microscopical preparations.     

Using wintergreen oil for mounting mosquito larvae: a safer alternative to xylene

Permanent mounting of fourth instar mosquito larvae is essential for identifying Aedes spp. This procedure requires extensive exposure to xylene, a clearing agent in the mounting process. We investigated wintergreen oil as a substitute for xylene. Five hundred larvae were mounted on slides to evaluate shrinkage or expansion of specimens after clearing using xylene or wintergreen oil. We examined the ventral brush and siphonal hair tufts for species identification and for preservation of morphological characteristics after clearing specimens in xylene or wintergreen oil. Shrinkage of the length of whole larvae and width of the head, thorax and abdomen after mounting was significantly greater after clearing with xylene than with wintergreen oil. The length of the comb scale nearest the ventral brush was similar for both clearing agents. The clarity of the specimens after mounting was improved by clearing with wintergreen oil, but the integrity of the ventral brush and siphonal hair tufts were similar for both clearing agents.     

Alcoholic Thionin Counterstaining Following Golgi Dichromate-Silver Impregnation

Brains of cats that had been fixed 2 months or longer in 10% formalin were cut into 3-6 mm. slices and impregnated by Golgi's dichromate-silver procedure (6% dichromate solution, 4-6 days; 1.5% silver nitrate solution 2 days). Sections 100 µ thick were cut after embedding in low melting point paraffin. Three changes of xylene and three of absolute alcohol were followed by staining 3-5 minutes in a saturated solution of thionin in absolute alcohol. The sections were dipped quickly in absolute alcohol and cleared in xylene, then differentiation was effected by an equal-parts mixture of absolute alcohol and xylene. A final clearing in three changes of xylene and mounting in Permount completed the process. Counter-staining was most successful when applied to freshly cut sections.     

A new method for identification of cement lines in undecalcified, plastic embedded sections of bone

A gallocyanin method for demonstrating cement lines in thin, undecalcified sections of bone has been developed that is compatible with prestaining with osteochrome before plastic embedding. After sectioning at 5 microns on the Jung K heavy duty microtome, the sections are attached to a microslide using Haupt's adhesive mounting medium, placed on a slide warmer at 37 C until completely dry, and deplasticized in xylene at 45 C for 16-24 hr. Sections are stained with 0.15% gallocyanin-5% chrome alum solution for 30 min, followed by staining in buffered Villanueva blood stain for 1-1 1/2 hr, quickly dehydrated, differentiated in equal parts xylene and 100% ethanol, cleared, and mounted in Eukitt's medium. Reversal lines appear as thin, scalloped, blue or purple lines approximately 0.3 micron wide, and arrest lines as thick, homogeneous, straight or evenly curved, dark blue or purple lines approximately 2 microns wide. The method also demonstrates abnormal halo volumes around osteocytes, old and new bone matrix, osteoid seams, and the granular mineralization front at the osteoid-bone interface. It promises to be valuable in the study of age-related bone loss, osteoporosis, and metabolic bone disease.     

A New Method for Identification of Cement Lines in Undecalcified, Plastic Embedded Sections of Bone

A gallocyanin method for demonstrating cement lines in thin, undecalcified sections of bone has been developed that is compatible with prestaining with ostcochrome before plastic embedding. After sectioning at 5 pm on the Jung K heavy duty microtome, the sections are attached to a microslide using Haupt's adhesive mounting medium, placed on a slide warmer at 37 C until completely dry, and deplasticized in xylene at 45 C for 16-44 hr. Sections are stained with 0.15% gallocyanin-5% chrome alum solution for 30 min, followed by staining in buffered Villanueva blood stain for 1-1 1/2 hr, quickly dehydrated, differentiated in equal parts xylene and 100% ethanol, cleared, and mounted in Eukitt's medium. Reversal lines appear as thin, scalloped, blue or purple lines approximately 0.3 pm wide, and arrest lines as thick, homogeneous, straight or evenly curved, dark blue or purple lines approximately 2 pm wide. The method also demonstrates abnormal halo volumes around ostcocytes, old and new bone matrix, osteoid seams, and the granular mineralization front at the osteoid-bone interface. It promises to be valuable in the study of age-related bone loss, osteoporosis, and metabolic bone disease.     

Stain Technology a Bone Stain for Osteoid Seams in Fresh, Unembedded, Mineralized Bone

Fresh, ground, mineralized bone sections 75-100 μ thick are stained 90 minutes or 48 hours in the Bone Stain, a preparation containing fast green FCF, orange G, basic fuchsin, and azure II. Surface stain is then removed by grinding under running water. Sections are washed in 0.1% zephiran chloride (benzalkonium chloride) or in 0.01% mild soap and again washed in tap water, followed with distilled water. Sections are next differentiated in 0.01% acetic acid in 95% methanol, dehydrated in 95% ethanol and 100% ethanol, cleared in alcohol:xylene 1:1, 1:4, 1:9 and 2 changes of xylol, and then mounted permanently in Eukitt's mounting media.

Osteoid seams stain either green to jade green or red to dark red, incompletely mineralized bone red or orange yellow, and the zone of demarcation light green. The walls of lacunae, canaliculae, feathered bone, procedural artifacts and periosteocyte lacunar low-density versions stain red.

The method helps in the differential diagnosis of certain metabolic bone diseases in human biopsy and autopsy material.     

Rapid Preparation of Thick Sections of Fleshy Plant Tissues

Methods are described for permanent micro-slide preparations of soft, large-celled plant tissues such as ripe fruit. Thick sections (200-800 [t) cut on a sliding microtome are aspirated in an aqueous killing agent; after fixing and washing, the sections are dehydrated and cleared in an alcohol-xylene series. Infiltration with 20, 30, and 40% solutions of mountant prior to mounting the sections is necessary to avoid too abrupt changes in the cleared tissues. Several staining methods have been successfully used for different purposes. The final preparations showed nearly perfect preservation of intact cells and intercellular spaces in their 3-dimension-al structure.     

Techniques for Clearing Ovules for Studies of Megagametophyte and Early Postfertilization Development in Rhododendron

Using ovule clearing, more than 33,600 ovules of Rhododendron nuttallii T. W. Booth (Ericaceae) were examined for megagametophyte and early postfertilization stages at daily intervals from anthesis until 3 weeks after pollination. Pretreatments with amyloglucosidase to digest integumentary and gametophyte starch and Stockwell's bleach to remove tannins from the integumentary epidermis were necessary. Ovules were cleared by a combination or modifications of Heir's four-and-a-half or Stelly's hem-alum-methyl salicylate techniques and were observed using differential interference contrast optics. The method proved suitable for large-scale quantitative studies of ovule development and fertilization. A general protocol is suggested as a starting point for ovule clearing studies.     

Euparal as A Mounting Medium for Preserving Fluorescence of Aniline Blue in Plant Material

Keeping quality of aniline blue-stained plant material can be greatly enhanced by substituting Euparal for aqueous aniline blue as the mounting medium. Pollen tubes which have been stained for other purposes (e.g. nuclei and chromosomes) and mounted in resinous medium can be restained with aniline blue and fluorescence of pollen tubes can still be observed.     

A Durable Nissl Stain for Frozen and Paraffin Sections

Frozen sections, 25-50 /j. thick, of formalin-fixed nervous tissues are mounted following the Albrecht gelatin technic. Paraffin sections, 15 p., are deparaffinized and transferred to absolute ethanol. The slides are then coated with celloidin. Both frozen and paraffin sections subsequently follow the same steps: absolute ethanol-chloroform (equal parts) for at least 20 min, 95% ethanol, 70% ethanol (1-3 min), then rinsed in distilled water. Sections are stained in Cresylechtviolett (Chroma) 0.5% aqueous solution containing 4 drops of glacial acetic acid per 100 ml, rinsed in distilled water, agitated in 70% ethanol until excess stain leaves the slide, and rinsed in 95% ethanol. Sections are then dehydrated in absolute ethanol, followed by butanol, cleared in xylene, and enclosed in permount.     

Vanillin-hydrochloric acid as a histochemical test for tannin.

Condensed tannin (leucoanthocyanins and catechins) can be demonstrated in fresh plant sections with saturated alcoholic vanillin followed by addition of concentrated HCl. Bright red vanillin-tannin condensates are formed immediately. Preparations may then be made permanent by mounting in a 1:1 mixture of Hoyer's medium and concentrated HCl. Some fading and loss of tannin occurs. The phloroglucinol-HCl test for lignin can also be made permanent with this acidic mountant.     

Vanillin-Hydrochloric Acid as a Histochemical Test for Tannin

Condensed tannin (leucoanthocyanins and catechins) can be demonstrated in fresh plant sections with saturated alcoholic vanillin followed by addition of concentrated HC1. Bright red vanillin-tannin condensates are formed immediately. Preparations may then be made permanent by mounting in a 1:1 mixture of Hoyer's medium and concentrated HC1. Some fading and loss of tannin occurs. The phloroglucinol-HC1 test for lignin can also be made permanent with this acidic mountant.